J. Eukaryot. Microbiol., 53(4), 2006 pp. 232–245 r 2006 The Author(s) Journal compilation r 2006 by the International Society of Protistologists DOI: 10.1111/j.1550-7408.2006.00100.x
Bonamia perspora n. sp. (Haplosporidia), a Parasite of the Oyster Ostreola equestris, is the First Bonamia Species Known to Produce Spores RYAN B. CARNEGIE,a EUGENE M. BURRESON,a P. MIKE HINE,b NANCY A. STOKES,a CORINNE AUDEMARD,a MELANIE J. BISHOPc,1 and CHARLES H. PETERSONc Virginia Institute of Marine Science, College of William and Mary, Gloucester Point, Virginia 23062, Virginia, USA, b National Centre for Disease Investigation, Ministry of Agriculture and Forestry, Upper Hutt, New Zealand 6007, New Zealand, and c University of North Carolina Institute of Marine Sciences, Morehead City, North Carolina 28557, North Carolina, USA a
ABSTRACT. Examination of the oyster Ostreola equestris as a potential reservoir host for a species of Bonamia discovered in Crassostrea ariakensis in North Carolina (NC), USA, revealed a second novel Bonamia sp. Histopathology, electron microscopy, and molecular phylogenetic analysis support the designation of a new parasite species, Bonamia perspora n. sp., which is the first Bonamia species shown to produce a typical haplosporidian spore with an orifice and hinged operculum. Spores were confirmed to be from B. perspora by fluorescent in situ hybridization. Bonamia perspora was found at Morehead City and Wilmington, NC, with an overall prevalence of 1.4% (31/2,144). Uninucleate, plasmodial, and sporogonic stages occurred almost exclusively in connective tissues; uninucleate stages (2–6 mm) were rarely observed in hemocytes. Spores were 4.3–6.4 mm in length. Ultrastructurally, uninucleate, diplokaryotic, and plasmodial stages resembled those of other spore-forming haplosporidians, but few haplosporosomes were present, and plasmodia were small. Spore ornamentation consisted of spore wall-derived, thin, flat ribbons that emerged haphazardly around the spore, and which terminated in what appeared to be four-pronged caps. Number of ribbons per spore ranged from 15 to 30, and their length ranged from 1.0 to 3.4 mm. Parsimony analysis identified B. perspora as a sister species to Bonamia ostreae. Key Words. Electron microscopy, fluorescent in situ hybridization, haplosporidian, phylogenetics, small subunit ribosomal DNA, sporulation, ultrastructure.
P
ROTISTS in the phylum Haplosporidia parasitize members of at least eight invertebrate phyla in freshwater and marine environments worldwide (Burreson and Ford 2004), but haplosporidians in the genus Bonamia are known exclusively from oysters in euhaline to polyhaline coastal environments (Carnegie and Cochennec-Laureau 2004). Bonamia species have also been noteworthy, in a phylum of spore-forming protists with presumably complex life cycles, for their apparent direct transmissibility (Elston, Kent, and Wilkinson 1987) and lack of a spore stage. Their characteristic cell form is a small (2–3 mm), uninucleate ‘‘microcell’’ (Pichot et al. 1980). Bonamia ostreae parasitizes Ostrea edulis in California, Washington, and Maine, USA (Elston, Farley, and Kent 1986; Friedman and Perkins 1994; Friedman et al. 1989), as well as Atlantic coastal Europe (Pichot et al. 1980); Bonamia exitiosa infects Tiostrea chilensis in southern New Zealand (Hine, Cochennec-Laureau, and Berthe 2001); and Bonamia roughleyi parasitizes Saccostrea glomerata in southeastern Australia (Cochennec-Laureau et al. 2003; Farley, Wolf, and Elston 1988). As yet undescribed species were found in T. chilensis in Chile and Ostrea puelchana in Argentina in the 1990s (Campalans, Rojas, and Gonzalez 2000; Kroeck and Montes 2005), and in an experimental population of Crassostrea ariakensis in North Carolina, USA, in 2003 (Burreson et al. 2004). The oyster Ostreola equestris (Say, 1834) inhabits euhaline to polyhaline waters from North Carolina, USA, south to Argentina (Harry 1985). A population of this little-studied, non-commercial species inhabits waters around Beaufort Inlet, NC (Wells 1961), near the site in Bogue Sound of the bonamiasis epizootic in C. ariakensis (Burreson et al. 2004). Owing to its presence within an aquaculture system containing the Bonamia sp.-infected and dying C. ariakensis, O. equestris was evaluated as a possible source of, or potential reservoir for, the C. ariakensis-pathogenic Bonamia sp. Examination of O. equestris revealed not only the Corresponding Author: R. B. Carnegie, Virginia Institute of Marine Science, College of William and Mary, Route 1208 Greate Road, Gloucester Point, Virginia 23062, USA—Telephone number: 804684-7713; FAX number: 804-684-7796; e-mail:
[email protected] 1 Present address: Department of Environmental Sciences, University of Technology, Sydney, PO Box 123, Broadway, Gore Hill, NSW 2007, Australia.
C. ariakensis-pathogenic Bonamia sp., but a second, novel Bonamia species that is the first to show strong histopathological and ultrastructural characteristics of the phylum to which ultrastructural data suggested (Pichot et al. 1980), and molecular phylogenetic analyses confirmed (Carnegie et al. 2000), the genus belongs. Uncharacteristically for a Bonamia species, it displays the diagnostic character of more typical haplosporidians (Sprague 1979): an ornamented spore with an orifice covered by a hinged operculum. A description of this new species, Bonamia perspora n. sp., based on histological, molecular, and ultrastructural analyses, is the focus of this manuscript.
MATERIALS AND METHODS Sample collection and processing. Ostreola equestris were collected from two locations in North Carolina. Samples from Bogue Sound, NC, were dredged from just offshore of the North Carolina State Port, at Morehead City (34143 0 N, 76142 0 W), monthly from January 2004 to August 2005 (n 5 55–200/month). Additional samples were obtained from an intertidal flat in Masonboro Sound at Wilmington, NC (34111 0 N, 77151 0 W) on 11 February, 23 May, and 22 June 2005 (n 5 100–200). The shell heights of the largest and smallest (January–September 2004) or of all (October 2004–August 2005) oysters in each sample were measured, and each oyster’s left valve was removed. Oyster identity was confirmed by noting the presence of chomata, which are absent in co-occurring Crassostrea spp. (Harry 1985). Small ( 3–5 mm3) pieces of gill/mantle (January 2004–April 2005) or visceral mass and gill (May–August 2005) were removed from oysters for DNA extraction and subsequent molecular analyses. These were either preserved first in 95% ethanol or placed directly in lysis solution (QIAamp DNA Kit, QIAGEN, Valencia, CA). Hearts or gill fragments of oysters collected from Bogue Sound in June 2004, and visceral mass and gill from oysters from Bogue Sound in May–August 2005 and from Wilmington in May and June 2005, were fixed in glutaraldehyde (2.5% [v/v] in 0.22 mmfiltered artificial sea water at 30 psu and 4 1C) for transmission electron microscopy (TEM). In all samples, remaining tissues were fixed for standard histopathology in Davidson’s fixative (Shaw and Battle 1957). Morphometric analysis and capture of
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light microscopic images were performed on an Olympus Provis light microscope outfitted with an Olympus DP70 digital camera. DNA extraction and Bonamia-generic PCR. DNA was extracted from each oyster tissue sample using a QIAamp DNA Kit (QIAGEN) and quantified on a GeneQuant pro spectrophotometer (Amersham Biosciences, Piscataway, NJ). Bonamia spp. small subunit (SSU) rDNA was detected by polymerase chain reaction (PCR) amplification using a modified version of the Carnegie et al. (2000) protocol. A 25-ml total reaction vol. included 1 PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs, bovine serum albumin at 0.4 mg/ml, primers (CF and CR; Carnegie et al. 2000) at 0.25 mM, AmpliTaq polymerase (Applied Biosystems, Foster City, CA) at 0.024 U/ml, and 0.5–1.0 ml (5 200–250 ng) template DNA. A 4-min initial denaturation at 94 1C was followed by 35 cycles of denaturation at 94 1C, annealing at 59 1C, and extension at 72 1C for 1 min each, and then by a final extension at 72 1C for 10 min. Products were electrophoresed on agarose gels, poststained with ethidium bromide, and evaluated under UV light. Small subunit rDNA sequencing. PCR products from four infected oysters collected in January 2004 from the Morehead City port area were used to generate the B. perspora SSU rDNA sequence. Amplification products from triplicate PCR reactions were pooled and cloned into the plasmid vector pCR4-TOPO using the TOPO TA Cloning kit (Invitrogen Life Technologies, Carlsbad, CA) and the inserts were subjected to simultaneous bidirectional sequencing using the Thermo Sequenase kit (Amersham Biosciences) and M13 forward and reverse infrared-labeled primers (LI-COR, Lincoln, NE). Sequencing reactions were electrophoresed and detected on a LI-COR Model 4200L automated sequencer. Initial sequence data were generated from the Bonamia spp.generic PCR products (generated using primers CF and CR). From within this sequenced region, we chose several candidate primers that could be paired with universal SSU rDNA-specific primers to amplify parasite, but not host, DNA. Successful amplifications were achieved with primer pairs 16S-A1BON-745R for the 5 0 end and BON-1310F116S-B for the 3 0 end of the gene (Table 1). PCR reaction components and cycling conditions were the same as for the Bonamia spp.-generic PCR except that the annealing temperatures were 53 1C for 16S-A1BON-745R and 50 1C for BON-1310F116S-B. Three to four clones per oyster for each PCR product were sequenced; these were aligned using the MacVector 7.0 software program (Oxford Molecular) to generate the consensus sequence. Bonamia sp.-specific PCR. Separate, species-specific PCR assays were developed for the C. ariakensis-pathogenic Bonamia sp. (Burreson et al. 2004) and for B. perspora. Both species-specific assays were applied to all Bonamia-generic PCR-positive oysters. Both reactions were performed in 25-ml vol. that included
template DNA (5 200–250 ng) and 1 PCR buffer, 1.5 mM MgCl2, 0.2 mM dNTPs, 0.4 mg/ml bovine serum albumin, primers (Table 1) at 0.5 mM, and Taq polymerase at 0.024 U/ml. Temperature cycling in both reactions began with a 4-min initial denaturation at 94 1C, which was followed by 35 cycles of denaturation at 94 1C for 30 s, annealing at 62 1C (C. ariakensis-pathogenic Bonamia sp. reaction) or 60 1C (B. perspora) for 30 s, and extension at 72 1C for 1.5 min, followed finally by a 5-min extension at 72 1C. Products were again electrophoresed on agarose gels, poststained with ethidium bromide, and evaluated under UV light. In situ hybridization. A fluorescent in situ hybridization (FISH) assay was used to reveal the range of B. perspora cell forms and their organ and tissue tropisms. Tissue sections (6 mm) were placed on positively charged slides (Colorfrosts/Plus, Fisher Scientific), dried, and deparaffinized with xylene (3 2 min). Slides were then rehydrated through a descending isopropanol series (100%, 3 30 s; 80%, 50%, and 30%, each for 30 s) into tap water (1 min), equilibrated in phosphate-buffered saline (PBS, 1 min), and digested with Proteinase K (100 mg/ml in PBS, 10 min at 37 1C); washed in PBS plus 0.2% (w/v) glycine (5 min), and acetylated with acetic anhydride (5% [v/v] in 0.1 M triethanolamine-HCl [pH 8.0], 10 min at room temperature); and washed in PBS (5 min), equilibrated in 5 SET (750 mM NaCl, 6.4 mM EDTA, 100 mM Tris base; 5 min at room temperature), and flooded with hybridization buffer (5 SET, 0.02% [w/v] bovine serum albumin, 0.025% [w/v] SDS; 10 min at 42 1C). Hybridization buffer was then drained off and replaced with 25 ml of hybridization buffer containing the appropriate oligonucleotide(s) (Table 1), all of which were designed for B. perspora specificity and purchased from Invitrogen Life Technologies with 5 0 Oregon Green labels. Slides were covered with parafilm coverslips and incubated overnight at 42 1C in humid chambers. They were washed the next day with 0.2 SET (three times at 42 1C for 2.5 min total), air dried, mounted with glycerol-in-PBS medium, and covered with glass coverslips. They were evaluated on an Olympus Provis epifluorescence microscope equipped with a redgreen dual bandpass filter. A competitive control experiment to assess the specificity of B. perspora probe binding included four treatments: a no probe control (25 ml of hybridization buffer only); a standard experimental treatment (B. perspora-specific probes each at 10 ng/ml); a competitive negative control (B. perspora-specific probes each at 10 ng/ml, plus unlabeled versions of these probes at 200 ng/ml); and a competitive positive control (B. perspora-specific probes each at 10 ng/ml, plus an unlabeled, non-specific oligonucleotide [a probe for a fungal parasite of deep-sea Bathymodiolus brevior in Fiji Basin] at 800 ng/ml). All but the no probe control were run in duplicate. Pre- and post-hybridization processing and microscopic evaluation were as above.
Table 1. Sequences of PCR primers and in situ hybridization probes used in this study. Primer/probe Bonamia CF Bonamia CR BON-745R BON-1310F 16S-A 16S-B OeBon-154F OeBon-472R B. perspora-ISH-1 B. perspora-ISH-2 B. perspora-ISH-3 B. perspora-ISH-4
Sequence
Use
Reference
5 0 -CGGGGGCATAATTCAGGAAC-3 0 5 0 -CCATCTGCTGGAGACACAG-3 0 5 0 -CTAATGCATTCAGGCGCGAG-3 0 5 0 -GAGACCCCACCCATCTAAC-3 0 5 0 -AACCTGGTTGATCCTGCCAGT-3 0 5 0 -GATCCTTCCGCAGGTTCACCTAC-3 0 5 0 -CAAAACCCCCGGCCACGTTC-3 0 5 0 -CATTCCGAATAGGCAACCAATC-3 0 5 0 -CCCCCGAACGTGGCC-3 0 5 0 -TGTGCGGTCGCGGACG-3 0 5 0 -GGCGGCCACTCTGCAC-3 0 5 0 -GCCGCGCATAAAAGAGC-3 0
Bonamia-generic PCR Bonamia-generic PCR Bonamia-generic PCR, Sequencing Bonamia-generic PCR, Sequencing Universal SSU PCR, Sequencing Universal SSU PCR, Sequencing B. perspora-specific PCR B. perspora-specific PCR B. perspora-specific FISH B. perspora-specific FISH B. perspora-specific FISH B. perspora-specific FISH
Carnegie et al. (2000) Carnegie et al. (2000) This study This study Medlin et al. (1988) Medlin et al. (1988) This study This study This study This study This study This study
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Molecular phylogenetics. The B. perspora SSU rDNA sequence was CLUSTAL-aligned (in MacVector 7.0 [Oxford Molecular], using default settings: an open gap penalty of 10.0, an extend gap penalty of 5.0, and with transitions weighted) with SSU rDNA sequences from most haplosporidians thus far characterized: Bonamia ostreae (GenBank Accession number AF262995), Bonamia exitiosa (AF337563), the Bonamia sp. from C. ariakensis in the USA, Haplosporidium louisiana (U47851), Haplosporidium nelsoni (U19538), Haplosporidium costale (AF387122), Haplosporidium pickfordi (AY452724), Haplosporidium lusitanicum (AY449713), a Haplosporidium sp. from Ruditapes decussatus (AY435093), Minchinia teredinis (U20319), Minchinia tapetis (AY449710), Minchinia chitonis (AY449711), a Minchinia sp. from Cyrenoida floridana (AY449712), Urosporidium crescens (U47852), and a Urosporidium sp. from Stictodora lari (AY449714). Additionally, we included sequences from two protists identified by Reece et al. (2004) as basal to the described haplosporidians: parasites from Pandalus platyceros (AY449715) and Haliotis iris (AF492442). Parsimony jackknife analysis of CLUSTAL-aligned sequences was performed using PAUP version 4.0d81(Swofford 2002) with settings at defaults. Before this analysis, 5 0 and 3 0 sequence ends were trimmed such that sequence coverage was uniform across all species in the analysis. TEM. Glutaraldehyde-fixed samples were rinsed once and stored in 0.22 mm-filtered seawater (30 psu) at 4 1C until PCR and histopathology results revealed B. perspora-infected individuals. These selected tissues were then post-fixed with 1% (w/v) osmium tetroxide buffered with 0.1 M sodium cacodylate at pH 7.2, dehydrated through a graded ethanol series including en-bloc staining with 1% (w/v) uranyl acetate added to the 70% ethanol stage of dehydration, and embedded in Spurr’s resin. Ultrathin ( 90 nm) sections were cut on a Reichert–Jung Ultracut E ultramicrotome, mounted on carbon-stabilized formvar-coated onehole grids, stained with Reynold’s lead citrate, and examined on a Zeiss CEM 902 TEM. Scanning electron microscopy. A 2 mm-square piece of infected oyster tissue that had been fixed in 2.5% (v/v) glutaraldehyde and held in sterile seawater (30 psu) at 4 1C was sonicated with a microprobe in a 1.5-ml microcentrifuge tube containing sterile seawater (30 psu). Sonication consisted of three 1-s bursts. A suspension of the sonicated tissue was puddled on four separate 12-mm-round coverglasses that had been coated with poly-Llysine. The sonicated tissue was allowed to settle for 1 h in a moist chamber. Coverglasses were washed by placing them in sterile seawater (30 psu) in small snap-cap vials before dehydration in a graded ethanol series. Material adhering to the coverglasses was critical point dried in liquid CO2 and then the coverglasses were mounted on stubs and sputter coated with Au:Pd. Observations were made on a LEO 435VP scanning electron microscope. RESULTS PCR detection. Evaluation of 1,644 O. equestris from the North Carolina State Port area of Bogue Sound, at Morehead City, between January 2004 and August 2005, revealed 20 oysters PCR-positive for B. perspora. The parasite was detected in 8/20 monthly samples, but never at a PCR prevalence greater than 5.6%. Bonamia perspora was also detected at low PCR prevalence in samples from Wilmington, NC, in May (4/200, 2.0%) and June (7/200, 3.5%) 2005. The February 2005 sample from Wilmington, NC, revealed no infections (Table 2). Histopathology. Subsequent histopathological examination of 26 PCR-positive oysters from both Bogue Sound and Wilmington confirmed the presence of uninucleate B. perspora cells in at least
Table 2. Bonamia perspora n. sp. infection status of Ostreola equestris samples from Morehead City and Wilmington, NC. Date
1/9/04 2/2/04 3/1/04 4/2/04 4/29/04 6/1/04 7/6/04 8/2/04 8/31/04 10/6/04 11/1/04 12/2/04 1/5/05 2/8/05 2/11/05 3/7/05 4/6/05 5/2/05 5/23/05 6/14/05 6/22/05 7/12/05 8/1/05
Location
Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Morehead City Wilmington Morehead City Morehead City Morehead City Wilmington Morehead City Wilmington Morehead City Morehead City
B. perspora-specific PCR
Histology results
# positive/ # examined (%)
# positive/ # examined
5/96 (5.2%) 1/64 (1.6%) 1/60 (1.7%) 0/60 3/60 (5.0%) 5/90 (5.6%) 0/60 0/59 0/60 0/60 0/60 0/60 0/60 0/60 0/100 0/60 0/55 2/200 (1.0%) 4/200 (2.0%) 0/120 7/200 (3.5%) 2/150 (1.3%) 1/150 (0.7%)
3/5 1/1 1/1 0/2 1/1
2/2 4/4 7/7 2/2 1/1
22 oysters. Bonamia perspora was not visually observed in four individuals. Uninucleate microcells (2.4–5.6 mm in diam.; mean SD: 4.0 0.7 mm; n 5 141; Fig. 1) with generally central to slightly eccentric nuclei were distributed throughout the connective tissues of infected animals, and were often abundant at the base of epithelia of the gut and hemolymph sinus. They seldom occurred inside hemocytes or any other host cells, and never occurred at the very high intensities typical of infection by other Bonamia spp. Diplokaryotic binucleate parasite cells co-occurred with uninucleate cell forms (Fig. 1), but at a lower abundance. These were sometimes spherical but more often ovoid, and measured 4.8 0.6 mm (mean SD; n 5 74) in the long dimension. Small plasmodia (3.7–9.7 mm in diam.; mean SD: 6.5 1.5 mm; n 5 45; Fig. 1, 2) were abundant in two B. perspora microcell-infected oysters from Morehead City in January 2004, and in one from February 2004. Like uninucleate and binucleate B. perspora cells, plasmodia were extracellular and distributed throughout oyster connective tissues. One Morehead City oyster from March 2004 displayed, along with uninucleate and binucleate B. perspora cells, greater numbers of larger plasmodia (6.8–17.8 mm in diam.; mean SD: 10.9 3.1 mm [n 5 30]), and a few packets of operculate spores in connective tissues within the visceral mass. Spores were 5.3 0.4 3.4 0.3 mm in size (mean SD; n 5 15). Another B. perspora-infected oyster from Morehead City in May 2005 also displayed some larger plasmodia (9.1–16.3 mm in diam.; mean SD: 13.0 2.4 mm; n 5 12), as well as numerous early sporonts. A full range of sporogonic cell forms (Fig. 2–6) was first observed in oysters sampled from Wilmington in June 2005. Of seven histologically confirmed B. perspora-infected oysters in this sample, four displayed sporonts and sporocysts in various stages of development: two in early sporogony (Fig. 2), one in midsporogony (Fig. 3), and one in late sporogony (Fig. 4), the last displaying thousands of free spores released within oyster tissues. In each case, sporonts or sporocysts (or released spores) were
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Fig. 1–4. Light microscopy of Bonamia perspora n. sp. in H & E-stained Ostreola equestris sections. All scale bars 5 10 mm. 1. Early infection, with expression of uninucleate (large arrows), binucleate (small arrow), and early plasmodial (arrowhead) B. perspora cell forms. 2. Early sporulation, with early (large arrow) and later () plasmodia, and early (small arrow) and later (arrowhead) sporonts, appearing in progression from the lower left to the upper right of the image along the base of the gut wall. 3. Later sporulation, with several small sporocyts (arrows) presented. 4. Numerous spores (arrows) released to the center of a disintegrating digestive tubule.
distributed throughout, and sometimes completely filled, oyster connective tissues. Sporonts and sporocysts measured 11.0– 26.4 mm in diam. (mean SD: 16.1 3.2 mm; n 5 60). Free spores were 5.0 0.4 4.0 0.3 mm in size (mean SD; n 5 31). Length ranged from 4.3 to 6.4 mm, width from 2.8 to 4.6 mm. The individual with late sporulation stages displayed widespread disintegration of digestive diverticula, while parasite cells (sporocysts and free spores) occurred in the lumens of many digestive tubules. Finally, an August 2005 oyster from Morehead City displayed small plasmodia along with uninucleate and binucleate B. perspora cells. Responses of the oyster hosts to B. perspora infection were variable. Hemocyte infiltration into infected tissues was generally strongest when infections were dominated by smaller parasite forms—uninucleate and binucleate B. perspora cells. Infiltration was generally weakest in hosts displaying plasmodial infections or parasite sporulation. In situ hybridization. Results of the competitive control FISH experiment (Fig. 5–9) supported binding of B. perspora probes to specific target sites. Relative to background tissue autofluorescence illustrated by the no probe control (Fig. 5), green fluorescence indicative of binding of B. perspora-specific probes was clear and unambiguous (Fig. 6). Specific probe binding was abolished by addition to the hybridization cocktail of a 20 concentration of unlabeled probes (Fig. 7), but not by addition of the same concentration of another oligo, an unlabeled probe specific for a fungal parasite of B. brevior (Fig. 8). Probes specific for
B. perspora did not hybridize to sections of oysters heavily infected with the C. ariakensis-pathogenic Bonamia sp. (Fig. 9). Application of this FISH assay to samples from Morehead City and Wilmington (Fig. 10–22) generated a staining pattern consistent with hybridization to all parasite stages observed histologically, including spores. Results suggested initial invasion of tissues by uninucleate and binucleate forms (Fig. 10), the former of which closely resembled B. ostreae cells evaluated using FISH (Carnegie, Barber, and Distel 2003): small green rings (the cell cytoplasm, containing the target SSU rRNA) enclosing a dark shadow (the nucleus) (Fig. 11). Uninucleate and binucleate cells then presumably developed into small plasmodia (Fig. 12) displaying distinct nuclei (Fig. 13). Plasmodia grew larger (Fig. 14), with nuclei becoming obscure and the cytoplasmic distribution of the ribosomes becoming more heterogeneous (Fig. 15). As these large B. perspora cells began sporulation (Fig. 16), a regular pattern of internal shadows again appeared within the B. perspora sporonts/early sporocysts (Fig. 17), indicative now of the position of developing sporoblasts. With advancing development (Fig. 18), sporoblasts increased in size and the surrounding cytoplasm became more limited (Fig. 19). As sporulation neared completion (Fig. 20), released spores filled connective tissues and were sometimes found in the lumens of disintegrating digestive tubules. Remaining late-stage sporocysts showed extreme reduction of cytoplasm surrounding spores, which were each enclosed now by intensely autofluorescent (red) spore walls (Fig. 21). Hybridization to the sporoplasm within individual spores occurred
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Fig. 5–9. Fluorescent in situ hybridization (FISH) controls for evaluation of specificity of Bonamia perspora n. sp.-specific probe binding. All scale bars 5 50 mm. 5. No probe control. 6. Experimental B. perspora-specific probe treatment, with parasite-specific probes at 10 ng/ml. 7. Competitive negative control, with specific probe hybridization abolished by addition of a 20 concentration (200 ng/ml) of unlabeled probes. 8. Competitive positive control, with specific probe hybridization unimpacted by addition of a 20 concentration of a haphazardly chosen oligonucleotide not specific for either Ostreola equestris or B. perspora n. sp. 9. Failure of hybridization to a Crassostrea ariakensis-pathogenic Bonamia sp. at high intensity in a C. ariakensis section.
frequently (Fig. 22). The failure of the probes to hybridize to some spores (Fig. 20, 21) was probably because of the impenetrability of unsectioned spores. Molecular phylogenetics. Parsimony analysis supported the monophyly of the Bonamia spp., and found 100% jackknife support for the placement of B. perspora in this clade (Fig. 23). Results suggest that B. perspora is a sister species to B. ostreae. Electron microscopy. Uninucleate, diplokaryotic binucleate, plasmodial, sporont, sporoblast, pre-spore and spore stages were recognized. Dimensions and parameters are given in Table 3. Uninucleate cells (Fig. 24). These cells possess a central spherical, sometimes irregular, dense nucleus with a nucleolus, large mitochondria with a lucent content and tubular cristae, a few haplosporosomes, lipid droplets, a few short to moderately long sections of smooth endoplasmic reticulum (sER), and dense ribosomes. Nuclear membrane-bound Golgi (NM-BG) are associated with haplosporogenesis. Binucleate cell (Fig. 25). The nuclei in a diplokaryon have 50 nm between the apposed nuclei with no internuclear chamber. Mitochondria surrounding the nuclei are more dense than in the uninucleate stage, and elongated with rounded cristae and dark matrix in some mitochondrial membranes. A few haplosporosomes, rare lipid droplets, several short or long parallel sections of sER and NM-BG, and dense ribosomes are present. Multinucleate plasmodia (Fig. 26). These were rare and small, containing o5 usually spherical nuclei/section, occasionally in diplokarya, with haplosporosomes, dense ring-shaped
mitochondria with rounded cristae, occasional NM-BG, no lipid droplets, short-long sER around the nuclei, and dense ribosomes (Table 3). Sporonts (Fig. 27–31). This stage was initially similar to plasmodia, but without haplosporosomes and with a thickened, folded plasma membrane (Fig. 27). There were one to four spherical nuclei/section lacking nucleoli, many lipid droplets, and circular dense or elongated mitochondria closely surrounding each nucleus (Fig. 28). A membrane develops separating the future sporoblasts from the folded thickened plasma membrane (Fig. 28). One to three NM-BG (Fig. 29) or rarely a reticulated structure resembling a spherule (Fig. 30) with a dense matrix suggest active synthesis (Fig. 29). Multiple membranes form in the cytoplasm of sporonts at the time of NM-BG activity, and then join to form uninucleate or binucleate sporoblasts (Fig. 29). Tubular structures extend from the surface of the sporont (Fig. 31). There are usually short, but occasionally long, parallel sections of sER, and sparse ribosomes. Later stages have one to seven nuclei/section, many lipid droplets, but fewer mitochondria, which are spherical and dense with rounded cristae (SDRC mitochondria). Sporoblasts (Fig. 32–34). Sporoblasts, lying within a sporocyst, are formed when membranes appear in the sporont cytoplasm to isolate single and paired nuclei. Partitioning of the sporont cytoplasm by these membranes marks the transition from sporont to sporocyst. Sporoblasts within the plasma membrane of the sporocyst initially have one to two spherical nuclei/sporoblast, NM-BG, no haplosporosomes, few lipid droplets, sparse
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Fig. 10–22. Progression and distribution throughout connective tissues of cell forms of Bonamia perspora n. sp. evaluated using fluorescent in situ hybridization (FISH). 10. Connective tissue colonization by primarily uninucleate and binucleate B. perspora cells. Scale bar 5 50 mm. 11. Two uninucleate B. perspora cells at higher magnification. Scale bar 5 5 mm. 12. Early plasmodia. Scale bar 5 50 mm. 13. Early plasmodium at higher magnification. Note distinct nuclei. Scale bar 5 5 mm. 14. Later plasmodia. Scale bar 5 50 mm. 15. Later plasmodia at higher magnification. Note heterogeneity of cytoplasm and absence of clearly resolved nuclei. Scale bar 5 5 mm. 16. Sporulation beginning, with early sporonts abundant. Scale bar 5 50 mm. 17. Early sporont at higher magnification. Scale bar 5 10 mm. 18. Late sporonts/early sporocysts. Scale bar 5 50 mm. 19. Several late sporonts/early sporocysts at higher magnification. Scale bar 5 10 mm. 20. Late sporulation, with abundant late sporocysts and numerous spores free in oyster tissues. Scale bar 5 50 mm. 21. Late sporocyst at higher magnification. Scale bar 5 10 mm. 22. Free spore at higher magnification. Green fluorescence indicates hybridization of probes to sporoplasm. Scale bar 5 5 mm.
3.5 (3–4)
2.0 (2)
1.5 1.2a, 0.6 0.9b (0–4a, 0–5b) 2.6 1.2a, 2.2 1.6b (0–8a, 0–5b) 4.0 4.5 (0–25) 4.9 2.9 (1–19) a
b
Including episporoplasm. Excluding episporoplasm.
6.0 1.6 (4–8)
0
0.6 0.9 (0–2) 5.8 2.9 (1–9)
10.8 7.6 (0–19) 23.9 15.3 (3–53)
Spores
Too few 3.0–4.5 Too few 2.6–3.0 1 Too dense 3.0 1.0 (0–10 and 2–4 extruded) 4.4 1.3 3.0–6.7 3.5 1.1 2.3–6.6 1–2 1.8 0.3 0
Immature spores Sporonts
4.6 0.3 4.3–5.0 3.0 0.5 2.6–3.6 2–3 1.3 0.1 2.8 3.4 (0–7) 4.0 0.9 3.1–5.5 2.8 0.9 1.9–4.5 2 1.5 0.3 2.4 2.9 (0–6)
Plasmodium Diplokaryon Uninucleate
4.1 0.6 2.5–4.8 2.6 0.5 1.8–3.0 1 1.5 0.2 4.5 3.0 (1–7) 136 20 1.0 1.4 (0–3) 7.0 1.0 (6–8)
ribosomes, SDRC mitochondria initially tightly surrounding the nucleus, rare short sER, but very many unit membrane-bound cytoplasmic vesicles, which were 170 25 114 19 nm (range 140–219 93–153 nm) (Table 3; Fig. 32) in diam. In section there were 9.3 2.9 (range 4–14) sporoblasts in each sporocyst. In apparently later stages with few vesicles, anucleate cytoplasm appears to engulf uninucleate sporoblasts (Fig. 33) to form the anucleate episporoplasm and uninucleate sporoplasm within the sporocyst (Fig. 34). Immature spores (Fig. 35–38). The episporoplasm contains lipid droplets and SDRC mitochondria. Spore wall forms at nodes between the episporoplasm and the sporoplasm. Small stacks of short sER are present. The sporoplasm is uninucleate, or rarely binucleate (Fig. 35), with a spherical equatorial to posterior nucleus with NM-BG in 14% of spores. A large apical spherule is present, usually containing osmiophilic matrix (63%) from which large ovoid light vesicles (OLVs) are formed (Fig. 36). In these, haplosporosome-sized bodies condense (Fig. 37), and pass into the cytoplasm, probably acquiring an outer membrane as they do so, to form haplosporosomes, with an internal membrane 3–4 nm wide. Lipid droplets are common, some of which pass into the space between the sporoplasm and episporoplasm to form lipidlike membranes (Fig. 38). Rarely the contents of the OLVs and/or haplosporosomes also pass into the space between the sporoplasm and episporoplasm. There are fewer SDRC mitochondria than in the sporoblasts. Rarely, short sections of sER are present. The immature spore wall and operculum comprise an exterior light layer 110 nm wide and an internal dark layer 25 nm wide. Filaments 110–300 nm wide and with no visible sub-structure pass from the external light layer through the episporoplasm to outside the spore.
Table 3. Dimensions and parameters of the developmental stages of Bonamia perspora n. sp.
Fig. 23. Parsimony jackknife analysis of haplosporidian small subunit rDNA sequences, with numbers at nodes denoting percentages of 1,000 jackknife replicates. Support was strong (100%) for inclusion of Bonamia perspora n. sp. within a monophyletic Bonamia clade.
11.1 3.7 4.2–17.2 7.2 2.6 3.2–12.3 1–7 2.2 0.6 0
Sporoblasts
4.2 0.5a, 3.6 0.5b 3.3–5.5a, 2.4–4.5b 3.2 0.2a, 2.6 0.2b 2.8–3.7a, 2.3–3.0b 1 1.3 0.1 0.7 1.0 (condensing)
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Length (mm) Length range (mm) Width (mm) (mean SD) Width (mm) (range) No. nuclei Mean SD diameter nuclei (mm) Mean SD no. haplosporosomes (range) Size haplosporosomes (nm) (mean SD) Mean SD no. lipid droplets (range) Mean SD no. mitochondria (range)
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Fig. 24–27. Cell forms of Bonamia perspora n. sp. prominent from tissue colonization through initiation of sporogony. 24. Uninucleate stage showing the central nucleus, swollen mitochondria, sparse haplosporosomes, and few moderately long profiles of smooth endoplasmic reticulum. 25. Extracellular diplokaryon showing the closely apposed nuclei, lack of nucleoli, mitochondria with rounded cristae around the nuclei, anastomosing endoplasmic reticulum by the smaller nucleus, long profiles of smooth endoplasmic reticulum, and two haplosporosome-like bodies (arrows). Scale bar 5 1 mm. 26. Early plasmodium with two nuclei, haplosporosomes (arrowheads), some ring-shaped mitochondria, many short dense profiles of smooth endoplasmic reticulum, and possibly nuclear membrane-bound Golgi (arrow). Scale bar 5 1 mm. 27. Sporont showing thickened folded plasma membrane, a large number of mitochondria, absence of haplosporosomes, and long parallel arrays of smooth endoplasmic reticulum (arrow). Perinuclear Golgi-like arrays contain a dense matrix (arrowheads). Scale bar 5 2 mm.
Mature spores (Fig. 39). This stage has no episporoplasm, and the sporoplasm is often dense, obscuring organelles. Lipid droplets, OLVs, spherule, and round dense mitochondria with rounded
cristae are present. Haplosporosomes are present in the sporoplasm and space between sporoplasm and spore wall. The operculum comprises three layers: an outer dark 95 nm across, a
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central light 30 nm across, and an inner dark 40 nm across. The spore wall also has three layers: an outer dark 110 nm across, a
central light 20 nm across, and an inner dark 40 nm across. Exsporulation was sometimes observed. Spores observed by SEM (Fig. 40) ranged from 3.6 to 4.2 mm in length with a mean length of 3.9 mm (n 5 15). They ranged from 2.9 to 3.3 mm in width with a mean width of 3.0 mm (n 5 15). The opercular lid ranged in diam. from 2.0 to 2.5 mm with a mean diam. of 2.2 mm (n 5 20). Spore ornamentation consisted of spore wall-derived, thin, flat ribbons that emerged haphazardly around the spore. Ribbons were often twisted along their length. The exact number of ribbons on each spore could not be determined, but varied considerably from about 15 to about 30 ribbons per spore. Ribbons were widest ( 300 nm) and thinnest ( 50 nm) where they emerged from the spore wall and then tapered gradually in width (to 100 nm), but became slightly thicker ( 75 nm), toward the tip. The end of each ribbon had a definite structure that seemed to consist of a 4-pronged cap (Fig. 41), although the exact nature of the end structure was not determined with certainty. Ribbons ranged in length from 1.0 to 3.4 mm with a mean length of 1.9 mm (n 5 25). DISCUSSION Histopathological evaluation, FISH, and molecular phylogenetic inference together confirmed that B. perspora is a Bonamia species of unprecedented diversity in cell form. Besides producing the uninucleate, binucleate, and plasmodial stages possessed by other members of this genus (Brehe´lin et al. 1982; Hine 1991a; Pichot et al. 1980), B. perspora produces sporonts, sporocysts, and spores as well. Possession by B. perspora of an ornamented, operculate spore, characteristic of the haplosporidian genera Haplosporidium and Minchinia, confirms that placement of Bonamia in the Haplosporidia is correct. This parasite is without question, therefore, a Bonamia sp. that produces spores, the first such case reported. Discovery of a spore-producing Bonamia sp. solves a taxonomic incongruency that has persisted over 25 yr. Pichot et al. (1980) suggested an affinity of Bonamia spp. to the Haplosporidia in their description of B. ostreae, noting that haplosporidians and B. ostreae all possess haplosporosomes. Perkins (1987, 1988) also placed Bonamia spp. in the Haplosporidia, and cited the absence of cell-within-a-cell forms as evidence against inclusion in the Paramyxea, whose members also possess haplosporosomes. Molecular phylogenetic analysis supported this assignment (Carnegie et al. 2000). Still, assignment of the non-spore-forming genus Bonamia to the Haplosporidia, a taxon whose members are defined by the structure of their spores (Sprague 1979), was unsatisfactory. We now recognize Bonamia as a spore-forming genus, like other haplosporidian genera, but one that includes several species that may have abandoned production of spores. Bonamia perspora is less like previously identified Bonamia spp. and it is more like typical haplosporidians not only in spore production but also in its extracellular residence and its affinity for connective tissues. Bonamia spp. are typically considered agents of ‘‘haemocytic parasitosis’’ (Balouet, Poder, and Cahour 1983), which invade host hemocytes and then evade host cytocidal Fig. 28–31. Sporont ultrastructure of Bonamia perspora n. sp. 28. Late sporont showing the mitochondria arrayed tightly around the nuclei, and membranes beginning to develop in the cytoplasm (arrow). Note the membrane (arrowheads) separating the future sporoblasts from the dense folded plasma membrane. Scale bar 5 2 mm. 29. An early sporoblast with nuclear membrane-bound Golgi containing dense matrix (arrow), suggesting active synthesis. Scale bar 5 1 mm. 30. A reticulated structure in the cytoplasm of an early sporont. Note the similarity to the spherule (Fig. 36). Scale bar 5 0.2 mm. 31. Tubular extension from the surface of a late sporont. Scale bar 5 0.2 mm.
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Fig. 32–35. Early spore development of Bonamia perspora n. sp. 32. Sporoblasts showing high degree of vesiculation of the cytoplasm. Scale bar 5 2 mm. 33. Sporoblast showing the anucleate portion (a), starting to surround the nucleate portion (b). Scale bar 5 2 mm. 34. Transverse sections of early sporoplasm (b) surrounded by early episporoplasm (a). The inclusion in one sporoplasm (arrow) may be the early spherule. Scale bar 5 2 mm. 35. Forming spore with two nuclei (n1, n2), with dense ovoid light vesicles (o). Note smooth endoplasmic reticulum in episporoplasm (arrow). Scale bar 5 1 mm.
mechanisms, dividing and ultimately rupturing hemocytes before entering new hemocytes and beginning the cycle anew (Montes, Anadon, and Azevedo 1994). This cycle of hemocyte invasion, parasite cell proliferation, host cell destruction, and hemocyte cell reinvasion is fueled by infiltration of hemocytes into an infected area (Bucke 1988), and results in a rapid multiplication of parasite numbers. Bonamia perspora, on the other hand, was rarely observed in oyster hemocytes, a strong indication that it pursues a life strategy very different from other Bonamia spp. Its systemic invasion of connective tissues as a uninucleate or small plasmodial form and the largely synchronous sporulation observed in those tissues in the few animals examined, in fact, is reminiscent of H. costale (Andrews and Castagna 1978).
The uninucleate and binucleate stages of B. perspora resemble those of other Bonamia spp., but they are comparatively large, with fewer and smaller haplosporosomes, and more lipid droplets and mitochondria (Table 4). They resemble B. exitiosa (Hine 1992; Hine and Wesney 1992, 1994), B. ostreae (Cochennec-Laureau and PMH, unpubl. data), H. costale (Perkins 1969), H. nelsoni (Perkins 1968), H. louisiana (Perkins 1979), a haplosporidian infecting abalone (Hine et al. 2002), and U. crescens (Perkins 1971) in possession of NM-BG. Gene sequencing places these latter species as basal to many other haplosporidian species (Reece et al. 2004). In B. perspora spores, haplosporogenesis occurs by condensation in the OLVs of haplosporosome-like bodies, which pass into
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Fig. 36–39. Later sporogony of Bonamia perspora n. sp. 36. Longitudinal section of an immature spore showing the spherule (s), the posterior nucleus (n), ovoid light vesicles (o), dense mitochondria (m), and lipid membranes between the sporoplasm and the wall (arrow). Scale bar 5 1 mm. 37. Haplosporosome-like bodies (b) forming in an ovoid light vesicle. Scale bar 5 0.2 mm. 38. Transverse sections of immature spores, showing the filament originating from the outer spore wall and passing through the episporoplasm (arrow). Note lipid being extruded into the space between the sporoplasm and the wall (arrowhead). Scale bar 5 1 mm. 39. Mature spore with dense content, including mitochondria (m), lipid (l), ovoid light vesicles (o), and extrasporoplasmic haplosporosomes (arrows). Scale bar 5 0.5 mm.
the cytoplasm, probably acquiring an outer membrane, similar to formation from electron-dense bodies, formative bodies or vesicles in Minchinia spp. (Azevedo and Corral 1989; Ball 1980; Comps and Tige´ 1997; Hillman, Ford, and Haskin 1990; McGovern
and Burreson 1990) and H. louisiana (Marchand and Sprague 1979; Perkins 1975). This differs from haplosporosome formation in dense vesicles (DVs) in Haplosporidium sp. infecting Saccostrea cuccullata, in which an internal membrane develops in the
CARNEGIE ET AL.—A BONAMIA THAT PRODUCES SPORES
Fig. 40–41. Scanning electron microscopy of spores of Bonamia perspora n. sp. 40. View of entire spore showing opercular lid () and ribbonlike ornamentation (arrows). Scale bar 5 1 mm. 41. Distal ends of ornamentation from a different spore than that in Fig. 40, showing morphology of the terminal caps (arrows). Scale bar 5 200 nm.
DVs, rather than the condensed haplosporosome-like bodies passing into the cytoplasm (Hine and Thorne 2002). The OLVs of B. perspora lack the striations reported from M. chitonis (see Ball 1980) and H. louisiana (Marchand and Sprague 1979; Perkins
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1975). This form of haplosporogenesis in the spore has not been reported from other Haplosporidium spp. The engulfment of the uninucleate half of a sporoblast by the anucleate half has been reported from Urosporidium jiroveci (Ormie`res, Sprague, and Bartoli 1973), Haplosporidium spp. (Cahour, Poder, and Balouet 1980; Ciancio, Srippa, and Izzo 1999; La Haye, Holland, and McLean 1984; Marchand and Sprague 1979) and Minchinia dentali (Desportes and Nashed 1983), and probably occurs in all spore-forming haplosporidians. The ejection of haplosporosomes into the space between the sporoplasm and the spore wall resembles the ejection of haplosporosomes or dense bodies in M. dentali (Desportes and Nashed 1983) and Haplosporidium spp. (Azevedo 1984; Azevedo and Corral 1987; Azevedo, Corral, and Perkins 1985; Burreson 2001; Comps and Pichot 1991), and possibly serves to open the operculum (Ormie`res and de Puytorac 1968). Bonamia perspora has several features in common with Minchinia spp. These include condensation of haplosporosomes in OLVs in the spore, and exsporulation in the host (Azevedo and Corral 1989; Ball 1980; Hillman et al. 1990; Desportes and Nashed 1983). The parallel arrays of sER resemble those of B. exitiosa (see Hine and Wesney 1994), although similar arrays have been illustrated from spores of M. chitonis (Ball 1980). Unlike Minchinia spp., B. perspora has lipid in the spores and the nuclei cluster tightly around the nucleus in the late sporonts. Several Haplosporidium spp. do have lipid in their spores (Hine and Thorne 2002; La Haye et al. 1984; Marchand and Sprague 1979; Perkins 1968, 1969) and mitochondria that cluster tightly around the nuclei (Bache`re and Grizel 1983; Ciancio et al. 1999; La Haye et al. 1984, Marchand and Sprague 1979; Perkins 1969). The delimitation of the sporoblasts by internal membranes has been reported from H. armoricanum (Fig. 9 in Bache`re and Grizel 1983, Fig. 3 in Pichot 1986), H. ascidiarum (Fig. 2 in Ciancio et al. 1999), and in association with Golgi in H. lusitanicum (Fig. 13, 14, and 16 in Azevedo et al. 1985). Exsporulation in the host also occurs in H. louisiana (Rosenfield, Buchanan, and Chapman 1969), and H. comatulae (La Haye et al. 1984). The spore ornamentation of B. perspora observed with SEM is unique in the phylum Haplosporidia. No other haplosporidian species has numerous, relatively short, thin, flat ribbons projecting from the spore wall and terminating in a definite capped structure. The ornamentation of B. perspora is most similar to that of Haplosporidium edule (Azevedo, Conchas, and Montes 2003). That species also has short projections from the spore wall, haphazardly arranged, and with a terminal structure. The ornamentation in H. edule, however, is round, not flat, and the terminal structure consists of relatively large opposing lobes. In other respects the spores of H. edule are quite unique, especially the numerous transverse projections of the outer spore wall that appear as folds with SEM.
Table 4. A comparison of main features of the uninucleate, binucleate and plasmodial stages combined of Bonamia spp. Species
Bonamia perspora n. sp.
Bonamia exitiosa1
Bonamia ostreae2
Bonamia sp.1
Eastern U.S.A. Ostreola equestris 3.9 1.0 33 137 19 62 0.7 1.0 38
New Zealand Tiostrea chilensis 3.1 0.4 15 7 148 11 32 0.7 0.9 44
Europe, U.S.A. Ostrea edulis 2.1 0.4 75 175 21 21 0.3 0.6 26
Australia Ostrea angasi 2.8 0.4 10 4 156 15 41 0.5 0.8 30
Location Host species Size (mm) No. haplosporosomes Size haplosporosomes (nm) No. mitochondria Mean no. lipid droplets % with lipid droplets 1 2
Hine (unpublished data). Engelsma & Hine (unpublished data).
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The observation of sporulation in particular alters our view of Bonamia spp. While other haplosporidians, with the possible exception of H. pickfordi (Barrow 1965), are only indirectly transmissible among their known hosts, direct transmission has been demonstrated for B. ostreae (Elston et al. 1987) and is presumed to characterize other Bonamia spp. The presence of spores in B. perspora suggests that other Bonamia spp. may also produce spores, but perhaps only under certain conditions that have not yet been observed. Developing spores were observed in only 6/22 (27.3%) confirmed B. perspora infections, or in o0.2% of all O. equestris examined by our laboratory. It is also possible that regular sporulation and what may be construed as a more conventional haplosporidian life cycle reflect a Bonamia sp. having been observed in its natural host. While B. exitiosa has not been observed to sporulate, it does have the best developed and most complex life cycle of described Bonamia spp. (Hine 1991a, b). It also has the longest standing documented relationship with its host (Farley et al. 1988), and may be truly endemic in T. chilensis populations. At the other extreme is B. ostreae, which has the simplest life cycle and the most minimal diversity in the cell forms it regularly displays, and concerning which is the strongest evidence for recent introduction to its known host (Elston et al. 1986; Katkansky, Dahlstrom, and Warner 1969). While microcells infecting vesicular connective tissue cells in Ostreola conchaphila from Yaquina Bay, Oregon (Farley et al. 1988) have been interpreted by some as B. ostreae, rather than Mikrocytos mackini (Arzul et al. 2005), it is not even clear that the source of B. ostreae on the west coast of North America was an oyster. More complex life cycles and more diverse arrays of cell forms may be found to characterize other Bonamia spp., such as B. ostreae, when these parasites are finally observed in their natural hosts.
TAXONOMIC SUMMARY Phylum Haplosporidia Caullery & Mesnil, 1899 Bonamia perspora n. sp. Diagnosis. Uninucleate microcells (2–6 mm) with central to slightly eccentric nuclei, occurring within connective tissues. Haplosporosomes present but few in number (5 3 cell 1), and small (136 20 nm). Mitochondria (6 2 cell 1) and lipid droplets (1.0 1.4 cell 1) abundant. Binucleate cell forms similar but larger in size (to 6 mm), less numerous. Progression of larger cell forms sometimes including small (o8 mm) and large (to 16 mm) plasmodia, sporonts, and sporocysts, the latter to 26 mm, within connective tissues. Spores 4–6 mm in length, 3–5 mm in width, when present, in disintegrating sporocysts or free within connective tissues and digestive tubules. Hinged opercular lid 2–3 mm in diameter. Episporoplasm absent. Ornamentation wall-derived ribbons, 15–30 spore 1; 1–3 mm long, 300 nm wide by 50 nm thick, tapering in width but increasing in thickness with distance from the spore wall; ending in a regular, four-pronged cap. Type host. Ostreola equestris (Say, 1834), crested or horse oyster. Type locality. Masonboro Sound at Wilmington, NC, USA, 34111 0 N, 77151 0 W, intertidal. Other locality. Bogue Sound at Morehead City, NC, USA, 34143 0 N, 76142 0 W, 4–6 m depth. Ribosomal sequences. Deposited to GenBank Accession number DQ356000. Etymology. The specific epithet is the Latin adjective perspora, meaning ‘‘by means of a spore.’’ Material deposited. Reference materials are deposited at the National Parasite Collection, U.S. Department of Agriculture, Beltsville, MD.
ACKNOWLEDGMENTS We thank Jeremy Braddy, Michael Ulery, and Tracy Hutcherson from the University of North Carolina Institute of Marine Sciences in Morehead City, and Ami Wilbur and Troy Alphin from the University of North Carolina Wilmington, for providing oyster samples. Virginia Institute of Marine Science (VIMS) Shellfish Pathology Laboratory staff Rita Crockett, Susan Denny, Martin Wunderly, and Lauren Martin performed the oyster histopathology, and Kristina Hill much of the sample processing and PCR diagnostics. Joe Scott and Megan Ward, College of William and Mary, and Patrice Mason, VIMS, graciously hosted the electron microscopy component of this study at their facilities, and shared their expertise. Grant support was provided by Virginia Sea Grant and the NOAA Chesapeake Bay Office. The helpful suggestions of three anonymous reviewers are gratefully acknowledged. This is VIMS contribution number 2732.
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